Nanoscale Pipetting for Controlled Chemistry in Small Arrayed Water

Nanoscale Pipetting for Controlled. Chemistry in Small Arrayed Water. Droplets Using a Double-Barrel Pipet. Kit T. Rodolfa,† Andreas Bruckbauer,† ...
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NANO LETTERS

Nanoscale Pipetting for Controlled Chemistry in Small Arrayed Water Droplets Using a Double-Barrel Pipet

2006 Vol. 6, No. 2 252-257

Kit T. Rodolfa,† Andreas Bruckbauer,† Dejian Zhou,† Andrew I. Schevchuk,‡ Yuri E. Korchev,‡ and David Klenerman*,† Department of Chemistry, UniVersity of Cambridge, Lensfield Road, Cambridge, CB2 1EW, United Kingdom, and DiVision of Medicine, Imperial College London, Hammersmith Hospital Campus, Du Cane Road, London, W12 0NN, United Kingdom Received November 10, 2005; Revised Manuscript Received December 19, 2005

ABSTRACT We present a new methodology which provides for the miniaturization of one of the most common tools in use in chemistry and biology laboratories todaysthe micropipet. We have used glass-fabricated double-barrel nanopipets to controllably produce arrayed water droplets with volumes as small as a few attoliters under an organic layer. We have addressed individual droplets and added controlled amounts of either additional volume or reagents from one of the barrels of the pipet. We demonstrate that this method can be used for miniaturized cell-free protein expression.

With the increasing global focus on nanoscience and new developments in nanotechnology, much work has been focused on the miniaturization of traditional laboratory techniques, experiments, and assays.1-19 For instance, nanoscale analogues of common implements such as pens (in the case of dip pen nanolithography16,17 or nanopipet-based writing18,20-23) and tweezers (in the case of nanotweezers15) have been developed. Laboratory studies in chemistry and biochemistry generally consist of mixing reagents in controlled amounts in order to produce a desired product or study the mechanism of a reaction. Performing these experiments on the smaller scale afforded by nanotechnology not only provides the benefit of using smaller amounts of potentially expensive and difficult to obtain reagents but also opens the door to new, and more complex, assays and even studying new phenomena. Toward this goal of the miniaturization of the biochemistry laboratory, we present here a new technique that is directly analogous to the macroscopic practice of chemistry, in which micropipets are used to add and mix reagents in the confined volume of an Eppendorf tube. Using voltage pulses applied to a double-barrel nanopipet with 100 nm barrel diameter, we can, in a highly controllable fashion, produce arrays of surface-immobilized drops with volumes as small as a few attoliters or as large as several hundred femtoliters. We further demonstrate that we can reliably * Corresponding author. E-mail: [email protected]. † University of Cambridge. ‡ Imperial College London. 10.1021/nl052215i CCC: $33.50 Published on Web 01/07/2006

© 2006 American Chemical Society

deliver species directly into these droplets at varied concentrations without detectably increasing the droplet size, allowing us to perform chemical reactions and biochemical assays within the confined space of specific droplets. It has previously been shown that glass-fabricated nanopipets can be controlled over a surface in a conducting solution for scanning ion conductance microscopy (SICM), using the ion current that flows between an electrode in the pipet and another in the bath for distance feedback control.24-26 SICM-based nanopipet techniques can be readily adapted to allow for controlled deposition of biomolecules onto a conditioned surface.20-22 This work has recently been extended to provide two-component depositions from a single tip by making use of double-barrel pipets working in air with a voltage applied between the two barrels.23 Here we show that double-barrel pipets operating under SICM control in an organic medium can be used to controllably deposit water droplets on surfaces as well as add reagents to specific droplets without adding to their volume. Droplet Creation. Our experiment is based on the use of a double-barrel pipet, with both barrels filled with electrolyte, working under oil as shown in Figure 1. As we have previously demonstrated in air,23 wetting of the tip by the solution inside the pipet allows an ion current, IDC, to flow between the two barrels when a potential is applied between them. As the tip is brought near a surface, the ion current is reduced, providing a signal for distance feedback control. Generally our pipet tips are controlled 75-

Figure 1. Schematic of the apparatus. A potential of 1-2 V is set by the digital signal processor (DSP) between Ag/AgCl electrodes in the two barrels of a glass-fabricated nanopipet creating an ion current flow between them, IDC. To provide more robust feedback, the tip is modulated in the z-direction. The resulting modulated ion current, IMOD, is measured by an ammeter (A) and used for feedback control by the DSP. Voltage pulses of 10-150 V are applied with the dc power supply unit (DC PSU) and used to deposit water droplets on the surface. Deposition was performed on uncoated glass coverslips. Drops are imaged with the fluorescence microscope. The inset shows an SEM image of a gold-coated pipet tip (scale bar is 200 nm).

125 nm above the surface.23 To provide more robust distance control, the tip is modulated in the z-direction (usually with magnitude of (50 nm), producing a modulated ion current, IMOD.26 Droplet production is not observed under our feedback conditions (1-2 V tip potential) but can be induced by pulsing to 25-100 V for 10-500 ms (see Figure 2 and supplementary movies 1 and 2). The drops could be produced reliably. The images in Figure 2 are representative data from drop arrays produced by our apparatus. Figure 2A shows 9 drops imaged optically with a mean diameter of 4.9 ( 0.1 µm and volume 32 ( 2 fL (assuming a hemisphere27). Fluorescence imaging was used in parts B and C of Figure 2 because the droplets were too small to be detected with white light illumination. Deconvoluted with our optical resolution of 350 nm, the mean drop sizes are 620 ( 20 nm in diameter (full width at half-maximum, fwhm, of fluorescence data) and 62 ( 7 aL volume in Figure 2B and 230 ( 140 nm diameter (fwhm) and 3 ( 2 aL volume in Figure 2C. A major advantage of our method is the high degree of control over drop size as illustrated in parts D and E of Figure 2, where a highly linear dependence of drop volume on the width of the applied voltage pulse (Figure 2D) and the pulse potential (Figure 2E) is evident. The volumes presented here assume a hemisphere, and error bars are propagated uncertainties from sample variance of drop diameter (five measurements with the same pipet in each case). The drops were Nano Lett., Vol. 6, No. 2, 2006

stable once deposited on the surface of the glass coverslip, and no droplet motion was detectable. Interestingly, for droplet production to occur, using tips with barrels of different sizes was found to be essential (see the Supporting Information for details).30 Drops could be deposited onto a glass surface by retracting the tip immediately after the voltage pulse. To promote adhesion of the droplets to the surface rather than the tip, chlorotrimethylsilane was used to make the outside of the glass nanopipets hydrophobic. Furthermore, while drop creation was observed under different organic layers, mineral oil was used in the present experiment. Presaturation of the organic liquid with water was found to be crucial to drop longevity, by preventing the water droplet from slowly dissolving into the oil. An interesting phenomenon is seen in Figure 2E, which clearly shows a cutoff voltage for drop creation (here, around 40 V). Beyond this potential, drop creation occurs readily, and quickly. For instance, the data in Figure 2D indicate a growth rate of 2.5 fL/ms when pulsed to 100 V. Below the cutoff potential, however, drops are not created at all. This threshold behavior is similar to that observed by Chiu and co-workers in a microfluidic system6 although our experiment has a different geometry using a double-barrel pipet very close to the surface. It appears the forces produced by the applied voltage need to overcome the interfacial tension between the water and oil to initiate droplet formation. Once an initial drop is formed, it requires much less energy to increase its size, thus accounting for the rapid expansion of drops when pulsed to a sufficiently high potential. While there is a high degree of reliability for drop production from any given tip, the particular parameters needed to produce drops of a given size can vary quite widely between tips. For instance, we have produced tips with a very wide range of cutoff potentialssbetween 15 and 75 V. We believe this is a function of variation in the size and shape of the tips produced by the pipet puller. In practice, this means that one must optimize the drop production parameters with each nanopipet used (note, for instance, that each part of Figure 2, except 2A and 2B, is produced by a different tip). This variation would be eliminated if the tips could be nanofabricated to control the size of each barrel. Volume Addition to Droplets. Addressing a droplet with the pipet by applying subsequent voltage pulses results in controllable droplet growth, as seen in Figure 3. This consists of bringing a tip into control over the droplet (with a tip potential of 1-2 V), applying a voltage pulse (as with drop production discussed above), and quickly withdrawing the tip after the pulse. Pulses lasting 300 ms and of 30 V were applied each time the drop was addressed. The top inset in Figure 3 shows representative data (of N ) 7 experiments) of additions made to one drop, imaged optically. Notice here that, while the drop diameter does not grow in a linear fashion, the volume is approximately linear (assuming a hemisphere shape) within experimental error. The data in Figure 3 indicate that an approximately constant amount of liquid comes out of the pipet with each pulse. However the volume of the initial droplet is about 40% of the volume of 253

Figure 2. (A) Optical image of nine water drops produced with the nanopipet (30 V, 300 ms pulse). Mean diameter is 4.9 ( 0.1 µm. (B) Fluorescent image of nine droplets containing Alexa 488 produced with the same nanopipet as in (A) (30 V, 50 ms pulse). Mean diameter is 620 ( 20 nm. (C) Fluorescent image of four tiny droplets produced with a separate nanopipet from (A) and (B) (30 V, 500 ms pulse). Mean diameter is 230 ( 140 nm. (D) Linear fit to calculated drop volumes vs voltage pulse width at 100 V. (E) Linear fit to calculated drop volumes vs voltage pulse amplitude with 250 ms pulse width. The error bars in (D) and (E) are propagated uncertainties from sample variance of drop diameters (N ) 5 measurements) with the same pipet in each case. Inset shows the process of drop creation: The tip is brought into contact with the surface and the drop is delivered with a voltage pulse and left on the surface by quickly retracting the pipet. Scale bars are (A) 5 µm, (B) 3 µm, and (C) 3 µm.

the subsequent additions providing additional evidence that formation of the initial droplet requires a threshold to be overcome but not the latter additions. Addition of Reagents. If a drop is addressed without applying a voltage pulse (i.e., the tip potential is maintained at the control potential of 1-2 V throughout the addressing), we have found that reagents can readily be added to the drop without adding to its volume (the diameter remains constant within the limits of our optical resolution of 350 nm). Figure 4 shows representative data of adding Alexa 488 dye. The data in the graph (Figure 4A) were produced by subsequent 20-s additions of dye with the tip retracted between additions.31 Figure 4B shows a series of images as dye is added to a droplet (see also Supporting Information movie 3): at 5.2 s, the tip contacts the droplet and the addition begins. Addition continues for ca. 4 s, and the tip is retracted fully at 10.3 s. It takes an additional 22 s before the droplet fluorescence has photobleached completely to its initial level. For Figure 4C, drops were produced with varied dwell times between the voltage pulse and withdrawing the tip. Total deposition time of the drops was 10.5, 1.5, and 0.5 s (including 0.5 s pulse width). The inset shows the fluorescence image of these drops while the graph is a line scan through this image (peak intensities are linearly 254

Figure 3. Linear fit to the increase in drop volume with subsequent addressing (30 V, 300 ms pulses). Error bars are from sample variance (N ) 5), and their increases from point to point are understood simply as a result of the variation in each addition addressing of a drop (deposition number 0 corresponds to the initial deposited droplet). The top inset shows representative optical images of an initial drop and its increases in size with four subsequent addressing pulses (scale bar is 5 µm). The bottom inset shows the process of sequential addressing of a droplet.

distributed with deposition time with R2 ) 0.987). Here, the reagents are delivered directly into the droplet with no need Nano Lett., Vol. 6, No. 2, 2006

Figure 4. (A) Linear fit of delivery of Alexa 488 with subsequent additions. Alexa 488 was added to a droplet by subsequent addressing for 20 s each without additional high-voltage pulse (tip potential during addressing was -1.0 V used for SPM control). Error bars represent uncertainties propagated from the noise in the fluorescent data. (B) A series of fluorescent images showing a representative addition of the dye to a droplet: before addition, during the addition, immediately after the tip has retracted, and during photobleaching. Scale bar is 2.5 µm. (C) Fluorescence images of three drops containing different amounts of Alexa 488 (inset) and line scan through the drops. Total deposition times for the drops (from left to right) were 10.5, 1.5, and 0.5 s. The inset shows the process of addition of Alexa 488 to a droplet.

to place species in the surrounding solution. Moreover, these data indicate that we can use the double-barrel pipet to deliver reagents from the pipet into a surface-immobilized droplet without any detectable removal of reagents already in the drop. The pipet appears to behave as a one-way valve although further studies are needed to confirm this and to elucidate the forces acting on molecules at the tip of the double-barrel pipet. An additional advantage is that since the tip potential during the reagent delivery is the same as that used for control (and thus well below the cutoff potential for drop creation), no volume is added to the droplet being addressed. An advantage unique to the double-barrel pipet is the potential for filling the two barrels with different reagents and controlling which is delivered by switching the sign of the voltage between the barrels. We have previously shown that the dye molecules, as well as various biomolecules, will generally migrate away from one electrode and toward the other.32 When two molecules migrate in the same fashion, this means they can easily be delivered independently from a double-barrel pipet. Generally, we have seen little evidence for mixing of the species in the two barrels, indicating that two reagents could be delivered individually and their reaction studied in the drop. Here, we present a proof-ofconcept experiment using Alexa 488 (green) and Alexa 647 (red) dyes. Figure 5 illustrates independent addition of the red and green dyes. In Figure 5A, a drop is created initially filled with the green dye, and subsequent 40 s additions of Nano Lett., Vol. 6, No. 2, 2006

Figure 5. Two-color fluorescence images. (A) A series of six images showing several alternate additions of Alexa 488 or Alexa 647 dye to a single droplet: creation of a drop containing Alexa 488; addition of Alexa 647 for 40 s; photobleached in both colors; addition of Alexa 647 for 40 s; addition of Alexa 488 for 40 s; followed by another 40 s of addition of Alexa 488. (B) Three drops created with 40 s additions of Alexa 488 (right), Alexa 647 (left), and both dyes (center).

red and green dyes produce the color changes shown in the image. Note that in the third image, the droplet has been photobleached in order to allow for red dye to be added and detected independently. Also note that, as with the addition of Alexa 488 seen in Figure 4, the drop size remains constant throughout these additions. Figure 5B shows the independent deposition of the two dyes (from a single pipet) into separate drops. First, two green drops were created (with 40 s 255

Figure 6. Expression of green fluorescent protein (GFP). Each bar on the graph represents the average of data from two experiments. The images above are representative data. Image sizes are 4.1 µm for the GFP production, 8.3 µm for the -DNA control, and 5.8 µm for the -MET control. -DNA and -MET control data were collected at 0.5 h after drop production.

deposition times), followed by a red drop (40 s deposition), and finally addition of the red dye for 40 s into the central green drop. Biochemical Experiments. We performed two proof-ofconcept experiments to demonstrate the applicability of the nanopipet to biological systems. We first used the doublebarrel pipet to form droplets containing fluorescein diphosphate and then added alkaline phosphotase, an enzyme which cleaves the fluorescein diphosphate to produce a fluorescent product,29 to individual droplets. This led to increased fluorescence with time, as the substrate was cleaved by the added enzyme, demonstrating that we could perform enzymology in droplets (see Supporting Information for details). We then used a linked transcription:translation kit to produce green fluorescent protein (GFP) in cell-sized droplets. Gene expression in cell-sized volumes has been demonstrated before using lipid vesicles;11 however this work required the presence of transcription:translation reagents in the bulk solution as well as in the vesicles themselves. Here, reagents are only present in the droplets and not in the surrounding organic layer. Immediately upon addition of the transcription mixture to the translation master mix, a pipet was filled with the reaction solution and used to produce droplets (150 V, 500 ms pulses). Droplets were photobleached after production to eliminate any background fluorescence and incubated at room temperature and GFP production was monitored over time using the fluorescence microscope with 488 nm excitation. -DNA and -MET control experiments were performed using the same procedure in the absence of GFPcoding DNA or the amino acid methionine (MET), respectively, and measured after 0.5 h of incubation (thus making them comparable to the first data point given). Figure 6 presents the average data from two repeats of each experiment as well as representative images of the fluorescence in each case. After 2 h of incubation, 300 GFP molecules were produced at a concentration of 37 nM in one drop (13 fL) and 550 GFP molecules at 17 nM concentration were produced in another drop of larger volume (54 fL). The GFP experiment described above shows that the biological molecules in the transcription:translation kit are 256

still functional after droplet formation using a voltage pulse. The pulse amplitude (25-100 V) for droplet production is relatively large compared to our control potential; however it is an order of magnitude smaller than the voltages that Yogi and co-workers used to see droplet production from a capillary tube operating in air.7 It should also be noted that the pipet tip introduces a high resistance (generally ca. 200 MΩ) and produces only a small current even under such high voltages (tens of nanoamperes during control, increasing to a maximum of only 500 nA during drop creation pulses of under 0.5 s in length).The electrodes are also far away from the tip (about 5 cm) preventing any problems associated with damage due electrolysis. We have shown in previous studies, operating continuously at voltages of 0.5-2 V, 21,28,29 that DNA, antibodies, and enzyme substrate maintain their functionality. Here we have used short duration pulses of higher voltage and shown for the specific biomolecules in these studies that they also maintain functionality. Therefore we have no evidence of damage to the biological material in any of the experiments performed with the nanopipet to date, although further experiments will be required to prove this is a general observation. The data presented here establish a novel methodology for the miniaturization of chemical and biochemical investigations through creation of very small water droplets in an organic environment using a double-barrel nanopipet. We have demonstrated that drops can be created with volumes as small as a few attoliters to as large as several hundred femtoliters, with a high degree of control over this size by adjusting the width or amplitude of the voltage pulse, applied between the two pipet barrels, to create the droplet. Furthermore, the double-barrel nanopipets can be used to reliably manipulate the drops to add volume and, more significantly, reagents. Most importantly, we have demonstrated that different reagents can be independently delivered into specific droplets from the same pipet and extension to a larger number of barrels in one pipet should be possible. The delivery is achieved with no observable increase in drop size and can be directly addressed to individual droplets. This research opens the door to using the droplets as tiny reaction chambers for chemical or biochemical reactions, “attoliter chemistry”, in an arrayed, highly miniaturized, and parallel fashion. Acknowledgment. This work was funded by the BBSRC. Winston Churchill Foundation provided a studentship to K.T.R. We thank Victor Ostanin for help with instrumentation and electronics and Tim Craggs for providing the GFP Vector used in the gene expression experiment. Supporting Information Available: Experimental details, the results of the experiment with fluorescein diphosphate and alkaline phosphatase, and three movies showing droplet creation and adding reagents to a droplet. This material is available free of charge via the Internet at http:// pubs.acs.org. References (1) Velev, O. D.; Prevo, B. G.; Bhatt, K. H. Nature 2003, 426, 515516.

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(2) Meister, A.; Liley, M.; Brugger, J.; Pugin, R.; Heinzelmann, H. Appl. Phys. Lett. 2004, 85, 6260-6262. (3) Kotz, K. T.; Gu, Y.; Faris, G. W. J. Am. Chem. Soc. 2005, 127, 5736-5737. (4) Schwartz, J. A.; Vykoukal, J. V.; Gascoyne, P. R. C. Lab Chip 2004, 4, 11-17. (5) Jorgensen, L.; Kim, D. H.; Vermehren, C.; Bjerregaard, S.; Frokjaer, S. J. Pharm. Sci. 2004, 93, 2994-3003. (6) He, M. Y.; Kuo, J. S.; Chiu, D. T. Appl. Phys. Lett. 2005, 87, art. no.-031916. (7) Yogi, O.; Kawakami, T.; Yamauchi, M.; Ye, J. Y.; Ishikawa, M. Anal. Chem. 2001, 73, 1896-1902. (8) Fischer, A.; Franco, A.; Oberholzer, T. ChemBioChem 2002, 3, 409417. (9) Katsura, S.; Yamaguchi, A.; Inami, H.; Matsuura, S.; Hirano, K.; Mizuno, A. Electrophoresis 2001, 22, 289-293. (10) Stamou, D.; Duschl, C.; Delamarche, E.; Vogel, H. Angew. Chem., Int. Ed. 2003, 42, 5580-5583. (11) Nomura, S.; Tsumoto, K.; Hamada, T.; Akiyoshi, K.; Nakatani, Y.; Yoshikawa, K. ChemBioChem 2003, 4, 1172-1175. (12) Bolinger, P. Y.; Stamou, D.; Vogel, H. J. Am. Chem. Soc. 2004, 126, 8594-8595. (13) Karlsson, A.; Sott, K.; Markstrom, M.; Davidson, M.; Konkoli, Z.; Orwar, O. J. Phys. Chem. B 2005, 109, 1609-1617. (14) Chiu, D. T.; Wilson, C. F.; Ryttsen, F.; Stromberg, A.; Farre, C.; Karlsson, A.; Nordholm, S.; Gaggar, A.; et al. Science 1999, 283, 1892-1895. (15) Kim, P.; Lieber, C. M. Science 1999, 286, 2148-2150. (16) Ginger, D. S.; Zhang, H.; Mirkin, C. A. Angew. Chem., Int. Ed. 2004, 43, 30-45. (17) Lee, K. B.; Park, S. J.; Mirkin, C. A.; Smith, J. C.; Mrksich, M. Science 2002, 295, 1702-1705. (18) Ying, L. M.; Bruckbauer, A.; Zhou, D. J.; Gorelik, J.; Shevchuk, A.; Lab, M.; Korchev, Y.; Klenerman, D. Phys. Chem. Chem. Phys. 2005, 7, 2859-2866. (19) Taha, H.; Marks, R. S.; Gheber, L. A.; Rousso, I.; Newman, J.; Sukenik, C.; Lewis, A. Appl. Phys. Lett. 2003, 83, 1041-1043. (20) Bruckbauer, A.; Ying, L. M.; Rothery, A. M.; Zhou, D. J.; Shevchuk, A. I.; Abell, C.; Korchev, Y. E.; Klenerman, D. J. Am. Chem. Soc. 2002, 124, 8810-8811. (21) Bruckbauer, A.; Zhou, D. J.; Ying, L. M.; Korchev, Y. E.; Abell, C.; Klenerman, D. J. Am. Chem. Soc. 2003, 125, 9834-9839.

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(22) Bruckbauer, A.; Zhou, D. J.; Kang, D. J.; Korchev, Y. E.; Abell, C.; Klenerman, D. J. Am. Chem. Soc. 2004, 126, 6508-6509. (23) Rodolfa, K. T.; Bruckbauer, A.; Zhou, D.; Korchev, Y. E.; Klenerman, D. Angew. Chem., Int. Ed. 2005, 44, 6854-6859. (24) Hansma, P. K.; Drake, B.; Marti, O.; Gould, S. A. C.; Prater, C. B. Science 1989, 243, 641-643. (25) Korchev, Y. E.; Bashford, C. L.; Milovanovic, M.; Vodyanoy, I.; Lab, M. J. Biophys. J. 1997, 73, 653-658. (26) Shevchuk, A. I.; Gorelik, J.; Harding, S. E.; Lab, M. J.; Klenerman, D.; Korchev, Y. E. Biophys. J. 2001, 81, 1759-1764. (27) Evidence is provided for this assumption by the fact that the diameters are strongly distributed with a cube-root relationship. Further, diameters are observed to diminish as the microscope focus is moved up through a drop. We note, however, that the drops may be somewhat less than hemispherical depending on the contact angle with the surface, meaning our volumes may be an overestimation. More research is necessary in order to better determine the exact three-dimensional structure of the drops. (28) Ying, L. M.; Bruckbauer, A.; Rothery, A. M.; Korchev, Y. E.; Klenerman, D. Anal. Chem. 2002, 74, 1380-1385. (29) Bruckbauer, A.; Zhou, D. J.; Ying, L. M.; Abell, C.; Klenerman, D. Nano Lett. 2004, 4, 1859-1862. (30) This asymmetry between the barrels varies considerably between tips, with ratios of the radii of the two barrels measured in the range of 1.2-1.6. (31) Very little dye was added in the initial 20-s addition, which is likely due to depletion of the dye in the tip region during drop creations here, the control potential used was -1.0 V while the drop was created with a pulse to +100 V. During this negative-to-positive pulse, the tip will have been depleted of Alexa 488 in the barrel that was subsequently used for delivery. The first addition (at -1.0 V) immediately followed the pulse, and thus there was little dye in the tip to be delivered. While the tip was retracted for 1-2 min between additions, however, the dye had time to return to the tip and subsequent additions proceeded in a linear fashion. (32) The direction of this migration will depend on the molecule’s charge as well as how it is affected by the electrophoretic, dielectrophoretic, and electroosmotic forces present in the tip. The dye molecules are small and negatively charged and, therefore, tend to migrate toward the anode.

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